Mutagenesis Advance Access published online on February 6, 2007
Mutagenesis, doi:10.1093/mutage/gem001
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Induction of DNA strand breaks and oxidative stress in HeLa cells by ethanol is dependent on CYP2E1 expression
The School of Biosciences, The University of Birmingham, Edgbaston, Birmingham B15 2TT, UK 1AstraZeneca R&D, Charnwood, Loughborough, Leicestershire LE11 5RH, UK
Induction of cytochrome P4502E1 (CYP2E1) is considered to be an important mechanism by which ethanol can cause toxicity related to oxidative stress both in vivo and in vitro. In the current study, we used HeLa cells with doxycycline-regulated CYP2E1 expression to test the hypothesis that induction of CYP2E1 could lead to secondary DNA oxidation that could potentially contribute to the carcinogenicity of ethanol in vivo. Overexpression of CYP2E1 protein was not associated with oxidative stress per se as assessed by markers of lipid peroxidation (cis-parinaric acid oxidation), glutathione depletion and elevation of intracellular reactive oxygen species (dichlorofluoroscin oxidation) in the presence or absence of ethanol substrate (10 mM, 24 h). Furthermore, there was no evidence of elevation of frequency of DNA strand breaks as assessed by the comet assay. In contrast, however, after pre-incubation of cells with L-buthionine-(S,R)-sulphoximine (BSO, 10 µM) which caused a 75% reduction in intracellular reduced glutathione (GSH) levels, CYP2E1 expression resulted in oxidative stress as assessed by all of these markers and DNA strand breaks but only in the presence of ethanol (10 mM). No effect was observed under these conditions in control cells not expressing CYP2E1. Furthermore, these effects could be attenuated by co-incubation with 1-aminobenzotriazole (0.5 mM), a suicide inhibitor of P450 activity. In conclusion, in this in vitro model CYP2E1-mediated interaction with ethanol results in the intracellular oxidative stress and the formation of DNA strand breaks which are detectable in cells pre-sensitized by depletion of intracellular levels of GSH.
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Introduction |
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Overexpression of cytochrome P4502E1 (CYP2E1), the ethanol-inducible isoform of P450, is of direct importance to human health and has been associated with a range of diseases including diabetes (1
–3), alcoholic liver disease and cancer (4–7). A variety of mechanisms have been implicated in the mechanism of ethanol hepatotoxicity. For example, CYP2E1 plays an important role in the metabolism of ethanol to acetaldehyde (ethanal) and the 1-hydroxy ethyl radical (8,9).
In addition, the reduction of molecular oxygen by NAD(P)H to water by CYP2E1 is widely considered to be substantially uncoupled under certain physiological situations and this may result in the generation of intracellular reactive oxygen species (ROS) (10–15). Indeed, microsomes prepared from rodent livers with induced levels of CYP2E1 show a greatly increased consumption of molecular oxygen and a concomitant production of hydrogen peroxide and superoxide compared to control microsomes even in the absence of substrate (16). Therefore, it seems likely that ethanol toxicity may also be related to CYP2E1 induction and futile cycling of oxygen and a disturbance of intracellular redox homeostasis. Previous studies have demonstrated that overexpression of CYP2E1 both in vivo and in vitro is associated with several cellular markers of oxidative stress including products of lipid peroxidation (4-hydroxynonenal and malondialdehyde), perturbation of Mitogen Activated Protein kinase signalling pathways as well as decreased viability as a result of both necrosis and apoptosis (17–23). However, it is not clear if CYP2E1 induction could lead to secondary DNA strand breaks potentially contributing to carcinogenesis.
The aim of the current study was to further investigate the relationship between CYP2E1 expression and intracellular oxidative stress and genotoxicity in the presence and absence of ethanol. Depletion of intracellular reduced glutathione (GSH) was also utilized to increase the sensitivity of cells to such effects. DNA damage was measured using the comet assay. For these studies, we used a previously described in vitro cell culture model (24) in which overexpression of CYP2E1 is under the regulation of the ‘Tet Off’ promoter system.
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Materials and methods |
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Cell culture
Tetracycline-regulated HeLa 2E1 cells expressing rabbit CYP2E1 were a generous gift from Professor Dennis Koop and have been characterized previously (24
). Cells were routinely cultured in T25 flasks (Nunc) at 37°C in a humidified, 5% CO2 atmosphere and maintained as a monolayer in Dulbecco's Modified Eagle Medium supplemented with 10% foetal bovine serum, 2 mM L-glutamine, 100 U/ml penicillin, 0.1 mg/ml streptomycin, 0.4 mg/ml G418, 0.25 µg/ml puromycin and 1 µ/ml doxycycline. Cells were passaged twice weekly using a standard trypsin–ethylenediaminetetraacetic acid (EDTA) protocol. Prior to commencement of experiments, cells were subcultured into six-well plates (or 96-well plates for the 3-(4,5-Dimethylthiazol-2-yl)MTT-2,5-diphenyltetrazolium bromide assay) and repression of CYP2E1 expression lifted by removal of doxycycline from the media. To ensure complete removal of doxycycline, cells were washed five times with phosphate-buffered saline (PBS) (3 ml for six-well plates and 100 µl for 96-well plates). Cells were then grown in the presence and absence of doxycycline (1 µg/ml) for 72 h. This procedure has previously been demonstrated to result in optimal induction of CYP2E1 (24). Test compounds were added as appropriate for the final 24 h of incubation.
Preparation of whole-cell protein
Whole-cell protein was prepared from confluent monolayers (T25 flask). Briefly, the media was removed and cells washed in PBS. Cells were then scraped into 0.5 ml of RIPA buffer [50 mM Tris–HCl pH 7.4, 150 mM NaCl, 1 mM Na2EDTA, 1% Triton X-100, 1% sodium deoxycholate, 0.1% sodium dodecyl sulphate (SDS)] containing 10 µl/ml mammalian protease inhibitor cocktail (Sigma-Aldrich UK, Dorset, UK) and mixed by pipetting four to five times before incubating on ice for 30 min with occasional vortexing. Following centrifugation (13 000x g, 10 min, 4°C), the supernatant was frozen at –80°C and retained for western blotting analysis.
Western blotting analysis
Protein extracts (30 µg) were resolved on a 12.5% SDS–polyacrylamide gel, transferred to nitrocellulose and blocked overnight at 4°C in blocking buffer [Tris-buffered saline (TBS)-0.05% Tween 20, 10% foetal calf serum]. Membranes were incubated with polyclonal goat anti-CYP2E1 (1 : 500 dilution, Oxford Biomedical Research, Oxford, UK, clone PR 32) in blocking buffer for 1 h 30 min at room temperature. Following washing (3x 10 min, TBS-0.05% Tween 20) membranes were incubated with horseradish peroxidase-conjugated rabbit anti-goat secondary antibody (1 : 2000 dilution, Dako UK, Dako Holdings Ltd, Ely, UK, P-0449) in blocking buffer for 1 h at room temperature and washed (as above). Bands were detected using enhanced chemiluminescence detection (Amersham Pharmacia Biotech, GE Healthcare UK Ltd, Bucks, UK).
MTT assay
The MTT assay was used to determine mitochondrial function. Cells were grown to 90% confluence in a 96-well culture plate (Costar, Fischer Scientific UK, Loughborough, UK). Following treatment, cells were washed with PBS, media replaced with MTT (0.45 mg/ml) containing growth media and subsequently incubated at 37°C for 3 h. Media was aspirated before the addition of dimethyl sulphoxide (DMSO) to solubilize the blue formazan product. Culture plates were gently rocked for 1 h before the absorbance was determined at 530 ± 40 nm against a DMSO blank.
Comet assay
Following the appropriate treatment, cells were washed in cold PBS and gently scraped into fresh PBS (1 ml). Cells were centrifuged (200x g, 5 min) and pellets re-suspended in PBS (150 µl). An aliquot of re-suspended cells (15 µl) was placed into a sterile tube containing low melting point agarose (150 µl) and this cell suspension transferred to a glass microscope slide (150 µl per slide; BDH Laboratory Supplies, Poole Dorset, UK), pre-coated with 0.5% normal melting point agarose. Glass cover slips (BDH) were added and slides placed on a metal tray over ice for 10 min. Cover slips were removed and slides incubated for 1 h at 4°C in lysis buffer (2.5 M NaCl, 0.1 M Na2EDTA, 10 mM Tris base, 1% sodium N-lauryl sarcosinate, 10% DMSO and 1% Triton X-100). Following lysis, slides were transferred to a horizontal electrophoresis tank (Pharmacia Biotech, GE Healthcare UK Ltd, Bucks, UK) containing electrophoresis buffer (0.75 M NaOH and 1 mM Na2EDTA, adjusted to pH 12.6) and DNA allowed to unwind for 30 min. DNA was subjected to electrophoresis (25 V, 30 min) and slides neutralized by flooding (3x 5 min) with neutralization buffer (0.4 M Tris, adjusted to pH 7.5). Slides were subsequently stained with 50 µl ethidium bromide solution (20 µg/ml). The slides were examined at x320 magnification (x32/0.40 dry objective) using a fluorescence microscope (Zeiss Axiovert 10, Carl Zeiss Ltd, Welwyn Garden City, UK), fitted with a 515- to 560-nm excitation filter and a barrier filter of 590 nm. A video camera (Kinetic Pulnix TM-765, Kinetic Imaging Liverpool, UK) received the images, which were analysed using a personal computer-based image analysis system Komet 3.0 Europe (Kinetic Imaging Ltd., Liverpool, UK). Images of 100 randomly selected nuclei were analysed per slide.
Measurement of percent tail DNA (TD %) was chosen to assess the extent of DNA damage as this has been shown to suffer much less from inter-run variation than other comet parameters because it is independent of electrophoresis voltage and run-time (25). Median values of three separate experiments were analysed using analysis of variance and post hoc Student's t-test, as recommended by Duez et al (26).
Determination of total GSH levels
Intracellular GSH was measured using the fluorometric method first described by Hissin and Hilf (27) with the following modifications. After treatment, cells were washed in PBS and lysed in cold whole-cell lysis buffer (0.1% Triton X-100, 5 mM Na2EDTA and 100 mM NaH2PO4, adjusted to pH 8.0). Protein precipitation buffer (50% w/v trichloroacetic acid, 5 mM Na2EDTA and 100 mM NaH2PO4) was added to the cell lysates, before centrifugation at 4°C (13 000x g, 10 min). Supernatants (100 µl) were transferred into 3 ml polystyrene fluorescence cuvettes (Sarstedt Ltd, Leicester, UK) containing phosphate–EDTA buffer (5 mM Na2EDTA and 100 mM NaH2PO4, adjusted to pH 8.0) (1.8 ml). A GSH standard curve was also constructed using a freshly made GSH stock (0.1 mg/ml in ice-cold phosphate–EDTA buffer) by adding the appropriate volume (0–20 µl) to cuvettes containing phosphate–EDTA buffer (1.8 ml) and 5% trichloroacetic acid solution (100 µl). o-phthalaldehyde (OPT) solution (1 mg/ml OPT in 100% methanol) (100 µl) was added to cuvettes containing standards and samples, before been mixed by agitation. Fluorescence was measured using a fluorometer (Perkin Elmer LS 50B, Perkin Elmer UK Ltd, Bucks, UK) with excitation at 340 nm (slit width 2.5 nm) and emission at 420 nm (slit width 4.0 nm). Total GSH (nmol) was determined from the standard curve and normalized to mass of total cellular protein (mg).
Assessment of lipid peroxidation
Lipid peroxidation was assessed by measuring loss of cis-parinaric acid (PNA) fluorescence as described previously by Carini et al (28). Briefly, cells were incubated with 10 µM PNA (Molecular Probes, Invitrogen, UK) at 37°C for 30 min in the dark. The media was then removed and cells washed three times with 3 ml warm PBS to remove unincorporated dye. Cells were then treated as appropriate prior to analysis. After treatment, cells were scraped into 2 ml PBS using a rubber policeman. The suspension was added to a fluorescence cuvette and the emission fluorescence of wavelength 455 nm (slit width 5 nm) measured using an excitement wavelength of 312 nm (slit width 5 nm). A blank (unlabelled cells) was measured and subtracted from all readings.
Assessment of intracellular ROS
Intracellular ROS (predominantly H2O2) was assessed by measuring intracellular oxidation of dichlorofluorescin to the fluorescent dye dichlorofluorescein as described by Carini et al (28). Briefly, cells were incubated with 10 µM dichlorofluorescein diacetate (Molecular Probes) at 37°C for 30 min in the dark. The media was then removed and cells washed three times with 3 ml warm PBS to remove unincorporated dye. Cells were then treated as appropriate prior to analysis. After treatment, cells were scraped into 2 ml PBS using a rubber policeman. The suspension was added to a fluorescence cuvette and the emission fluorescence of wavelength 520 nm (slit width 5 nm) measured using excitement wavelength of 502 nm (slit width 5 nm). A blank (unlabelled cells) was measured and subtracted from all readings.
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Results |
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Validation of experimental model
Prior to commencement of the main study, we confirmed by western blotting that this cell line had tetracycline-regulated expression of CYP2E1 and this is shown in Figure 1. In addition, there was no evidence of cytotoxicity as assessed by the MTT assay under any of the experimental conditions investigated (data not shown).
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Relationship between CYP2E1 and markers of oxidative stress
Overexpression of CYP2E1 in this cell line resulted in a small but consistent 15% reduction in intracellular glutathione levels from 16.8 ± 1.2 to 14.5 ± 1.4 nmol GSH per mg total protein in the absence of ethanol substrate (Figure 2) but this was not statistically significant (P = 0.055). However, in the presence of ethanol (10 mM), a statistically significant 21% (P < 0.05) reduction in intracellular levels of GSH in CYP2E1-expressing cells (14.1 ± 1.9 nmol/mg) was observed compared to cells not expressing CYP2E1 (17.7 ± 1.6 nmol/mg) (Figure 2). Although oxidised Glutathione (GSSG) was not measured, other parameters of oxidative stress were measured (discussed below). Treatment with L-buthionine-(S,R)-sulphoximine (BSO) (10 µM, 24 h) resulted in a 75% decrease in intracellular levels of GSH that was not dependent on the expression of CYP2E1 (Figure 2). No further depletion was observed following incubation of cells with both substrate (ethanol 10 mM) and BSO (10 µM).
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Lipid peroxidation (PNA fluorescence values) in untreated control and CYP2E1-expressing cells were 135.8 ± 34.7 and 149.2 ± 21.7 units/mg total protein, respectively, and there was no statistically significant difference. No change in PNA fluorescence (indicative of lipid peroxidation) was observed in either control or CYP2E1-expressing cells in the presence of substrate (ethanol, 10 mM) or BSO (10 µM) (Figure 3). In contrast, co-incubation with both BSO and ethanol resulted in a statistically significant reduction (P < 0.05) in PNA fluorescence in cells expressing CYP2E1 compared to non-expressing cells (Figure 3). This loss of fluorescence was ablated by incubation with the suicide cytochrome P450 inhibitor 1-aminobenzotriazole (0.5 mM, Figure 3).
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Dichlorofluorescein fluorescence (indicative of intracellular H2O2) values in untreated control and CYP2E1-expressing cells were 46.4 ± 14.6 and 52.7 ± 6.3 units/mg total protein, respectively, and there was no statistically significant difference. No increase in dichlorofluorescein fluorescence was observed in either control or CYP2E1-expressing cells in the presence of substrate (ethanol, 10 mM) or BSO (10 µM) (Figure 4). In contrast, co-incubation with both BSO and ethanol resulted in a statistically significant increase (P < 0.05) in dichlorofluorescein fluorescence in cells expressing CYP2E1 compared to non-expressing cells (Figure 4). This could be ablated by incubation with the suicide cytochrome P450 inhibitor 1-aminobenzotriazole (0.5 mM, Figure 4).
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Relationship between CYP2E1 and DNA strand breaks
TD % in untreated control and CYP2E1-expressing cells were 6.28 ± 1.2% and 5.98 ± 1.7%, respectively, and were not statistically significantly different (Figure 5A). No increase in TD % was observed in either control or CYP2E1-expressing cells in the presence of substrate (ethanol, 10 mM) or BSO (10 µM) alone (Figure 5A). However, co-incubation with both BSO and ethanol resulted in a statistically significant increase (P < 0.01) in TD % in cells expressing CYP2E1 compared to non-expressing cells (Figure 5A). Interestingly, elevation of DNA strand breaks in CYP2E1-expressing cells following treatment with ethanol and BSO was almost completely ablated by co-incubation with 1-aminobenzotriazole (0.5 mM, Figure 5B). Titration with decreasing concentrations of doxycycline (1000–0 ng/ml) resulted in a concentration-dependent increase in DNA strand breaks as assessed by the comet assay (Figure 6) further indicating the CYP2E1 dependency.
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Discussion |
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The mitochondrial electron transport chain is widely considered to be the single major source of intracellular ROS. However, recent studies have also established that several isoforms of cytochrome P450 may under certain conditions contribute significantly to the intracellular burden of ROS reviewed by Zanger et al (29
). For example, work by Kinoshita et al (30) and Imaoka et al (31) has demonstrated that in rodent liver, phenobarbital-induced cytochrome P450 is a likely mediator of intracellular oxidative stress including oxidative damage to DNA. Induction of other isoforms has also been associated with oxidative stress, for example, CYP1A1 (32,33) and CYP1B1 (34). Interestingly, CYP1A1 gene expression is negatively regulated by intracellular oxidative stress (35,36) suggesting the possible existence of a cellular negative feedback loop to regulate CYP1A1-mediated generation of ROS. In addition, cytochrome P450s have also been implicated as a contributor to oxidative stress following ischaemic reperfusion injury such as myocardial infarction (37,38).
In the current study, we have demonstrated in an in vitro cell culture model that overexpression of CYP2E1 results in induction of DNA strand breaks (as assessed by the comet assay) but only under conditions of GSH depletion by co-incubation with a non-toxic (10 µM) dose of BSO, an inhibitor of GSH synthesis. This effect was almost entirely inhibited by 1-aminobenzotriazole strongly suggesting a direct role of CYP2E1. Our data further highlights the importance of GSH in the protection of cells from CYP2E1-derived reactive species and subsequent oxidative stress and is in agreement with the previous study of Wu and Cederbaum (21) who demonstrated the importance of GSH in the protection of cells overexpressing CYP2E1 from both apoptosis and necrosis in vitro. In contrast to our study where we used BSO (10 µM, 24 h) which caused a 75% reduction in intracellular GSH levels to sensitize cells to CYP2E1-dependent oxidative stress, previous studies used much higher concentrations of BSO (300 µM) that resulted in levels of GSH depletion that were directly toxic to cells even in the absence of CYP2E1 expression. In addition, these studies did not investigate the possible formation of DNA strand breaks under these experimental conditions.
Toxicity of CYP2E1 expression in cells in GSH-depleted cells has been related to activation of p38 MAP kinase and a reduction in nuclear factor 
B activity (23) and protein kinase C activation. Induction of the catalytic subunit of glutamate–cysteine ligase (GCLM) in HepG2 cells overexpressing CYP2E1 has also been demonstrated (39,40). Very recently, Gong and Cederbaum (41) have shown that this induction is a result of activation of the redox-sensitive transcription factor Nrf2 which activates transcription of genes with antioxidant response elements in their promoters including GCLM and haem oxygenase 1. This effect was observed both in rodent liver and HepG2 cells and is likely to represent a cellular adaptation to CYP2E1-mediated oxidative stress.
Interestingly, serum deprivation which is associated with oxidative stress also sensitizes cells to CYP2E1-related toxicity (42). This sensitivity could be ‘rescued’ by treatment of cells with antioxidants. In addition, studies with primary rat hepatocytes have suggested a possible role of cytochrome P450 in toxicity and oxidative damage in the presence of catalase and glutathione peroxidase inhibitors (43).
To our knowledge, the current study is the first to investigate the relationship between CYP2E1 overexpression, oxidative stress and DNA strand breaks in an in vitro cell culture model and how this might relate to DNA damage by ethanol. In vivo, previous studies (44,45) have demonstrated that chronic ethanol exposure is associated with DNA strand breaks in rodent liver. However, it is not clear if this is a direct effect or an indirect mechanism such as through an inflammatory response. Studies by Navasumrit et al (46) have demonstrated elevated levels of etheno adducts in the genomic DNA of liver from ethanol-treated rats suggesting that breakdown products of lipid peroxidation (malondialdehyde and 4-hydroxynonenal) are at least partially responsible for ethanol-mediated DNA damage. Previous studies in rodent liver also indicate an elevation in oxidative DNA damage (8-oxo dG) following ethanol treatment suggesting a role of ROS.
In the current study, the lack of DNA strand breaks observed in the absence of ethanol as a substrate suggests either that it is a CYP2E1-derived metabolite of ethanol that is responsible for the formation of DNA strand breaks or that production of ROS by futile cycling of CYP2E1 is enhanced by ethanol. In support of the former interpretation, it is known that CYP2E1 has the potential to metabolize ethanol to several electrophilic and potentially DNA damaging moieties including ethanal and the 1-hydroxy ethyl radical. However, it is not clear if the oxidative stress observed (and potentially associated DNA damage) is mediated by these metabolites or by reactive oxygen release from CYP2E1.
In our experimental model, at least, levels of ROS generation from CYP2E1 and ethanol were not sufficiently high to cause measurable DNA damage without prior sensitization by GSH depletion. In summary, both overexpression of CYP2E1 and ethanol exposure have been associated with cancer risk (4–7). In the current study, we have demonstrated that ethanol treatment in cells that overexpress of CYP2E1 results in formation of DNA strand breaks detectable when protective GSH is depleted. This may contribute to the mechanism of alcohol-induced hepatocarcinogenesis in vivo.
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Notes |
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* To whom correspondence should be addressed. Tel: +44 121 414 5906; Fax: +44 121 414 5925; Email: N.Hodges{at}bham.ac.uk
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Received on October 27, 2006; revised on December 14, 2006; accepted on December 18, 2006.
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